Propagation Methods


Photo of C. reginae culture tubePhoto of C. guttatum culture tubePhoto of C. formosanum culture tube

Protocorms and young seedlings of three Cypripedium species. From left to right: C. reginae, C. guttatum, and C. formosanum. In each case, the culture tube occupying the width of the photo is 2.5 cm (1 inch) across.


Introduction.

In this section I will present the methods we currently use to produce all of our Cypripedium seedlings.  Orchidists use one of two procedures in the lab to grow orchids from seed: 1) collecting still-developing seed from immature capsules ("green pod culture") or 2) beginning with fully mature seed.  The advantage to using immature seed from unripe capsules is that the seed is not dormant;  the embryo is actively growing and can be transferred directly to in vitro culture, and there is no problem overcoming dormancy because the seed is not yet dormant.  The main difficulty with this procedure is that the seed must be collected during a very short window of time in which the embryo can survive transfer from the mother plant and before the mother plant imposes dormancy on the seed.  Harvesting and culturing mature seed is much more lenient regarding the time of collection of the seed, but the main difficulty is that dormancy in the seed must be broken artificially.  Additional advantages of the use of mature seed are that 1) the mature seed can be stored for long periods of time and remain viable and 2) because the seed is dormant, it can be mailed or otherwise exchanged with other propagators worldwide.  At Spangle Creek Labs we propagate from mature seed almost exclusively.

Seed Collection.

Collecting mature seed is relatively easy, but timing is important.  The seed should be collected when it truly is mature and the mother plant has imparted all vital materials to the seed, but the seed capsule needs to be harvested before it dehisces, that is, splits open naturally.  The perfect time to collect the capsule would be a few seconds before dehiscence, but such precision is not necessary.  What is important is to collect the seed after the mother plant has given the seed everything she has to offer, but before the capsule splits.  If the capsule opens naturally, rain or dew may provide sufficient moisture that the seed becomes moldy, or worst of all, the seed falls out and becomes lost.  In moldy seed, fungal hyphae reach deep into the seed and its embryo, and surface sterilization in the lab cannot eliminate the fungus.  In practice, Cyp seed can be harvested within days or even weeks of the optimal time, but take care to avoid collecting too early or too late.

  

Treatment of Mature Seed after Collection.

After collection of a seed capsule, it should be opened in the lab with a couple slices down the length of the capsule with a sharp knife or scalpel.  The seed should be poured and scraped out onto a piece of clean paper for drying in the air.  The purpose of drying is to make sure that the seed does not retain sufficient moisture to allow the growth of mold.  Depending on the ambient relative humidity, the length of drying may vary from two to five or more days.  The drying period should not be extended excessively as Cyp seed loses vigour rapidly at room temperature.

After drying, I place the seed in small glass vials with plastic caps for temporary storage in the refrigerator at 0 to 4 degrees C until sowing.  The seed can be kept at these temperatures for several months without much deterioration in vitality.  I have had seed of some species stored up to three years in the refrigerator and still germinate, but lower temperatures are far better for long term storage.  Cyp seed is not harmed by freezing, and I store seed long term in a freezer with temperatures well below 0 C.

Mailing Seed.

As  I mentioned above, one of the main advantages of working with mature seed is that it can be exchanged by mail with growers all over the world.  A difficulty with such exchange, however, is that mail is often handled very roughly.  In the U.S., mail sorting machines are brutal and will crush seed unless it is protected by a strong container.  I once even had a sturdy glass vial of seed smashed by this equipment.  All too often I have  received orchid seed mailed in paper envelopes from well meaning donors only to see under the microscope that the seed had been crushed into a form similar to flakes of breakfast cereal.  Such seed will not germinate. Even seed mailed in envelopes padded with bubble plastic often arrives damaged to the point that there is minimal germination.  To my knowledge, a 1992 article by Warren Stoutamire was the first to alert orchid propagators to this problem.

The solution to this difficulty is to mail seed in some sort of sturdy, rigid container.  My preference is to use micro-centrifuge tubes made of very strong plastic, but anything strong enough to survive the mail-sorting machines will do.  Examples include old fashioned 35 mm plastic film canisters, compact disc boxes, and small jewelery boxes.  The essential thing is to prevent the seeds from being crushed.

Sowing Seed.

        Preliminary Treatment of Seed.

Cypripedium seed, like orchid seed in general, consists of an embryo within a seed coat known as a testa.  The testa is water repellent, and the seed has a large air space between the embryo and testa so the seed tends to float on water.  The seed, however, has a small opening at the micropyle end, and so with appropriate techniques, the air can be removed from the seed thus facilitating the entry of aqueous solutions to surround the embryo intimately.  These solutions are either chemicals whose purpose is to remove impediments to germination on/in the embryo, or nutrient solutions the embryo needs for germination and subsequent growth.

To remove air from within the testa, the seed is given a preliminary soak, under partial vacuum, in water purified by reverse osmosis (RO), a technique I first learned from an article by Allan Anderson (1989).  In my application, I place a smidgen of seed in a small, ~100 mL, glass bottle and add approximately 20 mL of RO water and a drop of the biological surfactant Tween20.  I then put the lid on the bottle and gently shake it for a couple minutes.  Next, I remove the lid from the bottle and replace it with a rubber stopper connected by plastic tubing to a hand vacuum pump.  This equipment is shown at the left.  In my experience the 
Bottle and Vacuum Pumpvacuum pump can be purchased least expensively as a component of a do-it-yourself automobile brake bleeding kit.  The pump shown is capable of reducing atmospheric pressure by roughly 600 mm Hg, but a little extra pumping is required every half hour or so as air gradually leaks back into the system.  Periodically, the bottle is returned to full atmospheric pressure and observed to see whether the seeds sink to the bottom.  Usually several cycles of pumping and re-pressuring are required to get most seeds to sink.  Presumably, the low density seeds that do not sink are defective and can be pipetted off along with most of the water. When most seeds have sunk, the overlying water is removed with a pipette along with any floating debris and seeds.  At this point, the seeds are ready for bleaching.  


Bleaching

Because the outside of mature orchid seeds from open capsules is infested with microorganisms, surface sterilization of the seed in chlorine bleach has long been a standard practice in orchid lab propagation.  There is, however, a second extremely important function of the bleaching of the seed:  Extended bleaching results in removal or destruction of dormancy-promoting factors and results in greatly enhanced germination.

That lengthy bleaching speeds germination and increases percentage germination was discovered serendipitously by Allan Anderson (1989) when he accidentally left Cypripedium seeds in the bleaching solution too long.  The beneficial effect of extended bleaching on the seeds of other genera of terrestrial orchids was found independently by Ben Lindén (1980) in Finland in a series of controlled experiments.  The mechanism for the enhancement of germination by lengthy bleaching, that is, bleaching longer than necessary merely for surface sterilization, is not definitely known.  The effect seems to be some combination of demolition of the hydrophobic carapace, the residual layer of the inner integument covering the embryo, by the bleach and the removal and/or destruction of germination inhibiting chemical compounds by the highly oxidizing bleach solution.  In much of the orchid propagation literature, the positive effect of bleaching on germination is attributed to action of the bleach on the testa or seed coat with claims that bleaching destroys the cutinaceous nature of the testa, thus allowing water or germination medium to enter the cavity of the seed containing the embryo.  I am convinced, however, that this is not the mechanism, for the testa has an opening at the micropylar end that permits entry of liquid into the space containing the embryo.  That the bleach readily enters the seed coat after the vacuum treatment and directly attacks the carapace can easily be observed under the microscope, for even in the early stages of bleaching the formation of gas bubbles can be seen in the seed cavity as the bleach reacts with the carapace.  Moreover, in many Cypripedium seeds the colored carapace can be seen to be removed as the bleaching process continues.  In summary, there are two reasons for bleaching: 1) to accomplish surface sterilization of the seed, and 2) to remove germination inhibitors.


The bleaching solution is prepared by diluting a commercial sodium hypochlorite (NaOCl) household bleach, namely Clorox®, with water to obtain a bleaching solution that is 0.5% NaOCl.  I do the dilution in a 100 mL graduated cylinder, diluting the commercial bleach with enough water to make 100 mL of solution.  I then pipette roughly 10 mL of this solution onto the seeds in the bottle, being very gentle so as not to cause any seeds to float again.  After this10 mL quantity of solution has been pipetted into the bottle, I gently pour the rest of the solution from the cylinder into the bottle and agitate the contents from time to time.

Bleaching times of 5-10 minutes are generally sufficient for surface sterilization unless the seeds are actually infected, but most Cyp species require considerably longer bleaching for removal of germination inhibitors. The optimal duration of the bleaching is usually in the range of 15-150 minutes and varies considerably not only among different species but within different batches of seed of the same species.  Within a species, different clones may have considerably different optimal bleaching times, and this time may vary from one year to the next for seed of the same clone.

In preparing the bleaching solution, use of fresh commercial bleach is extremely important, for the activity of the bleach decreases with time;  the shelf life is not very long.  I try to purchase fresh bleach at a large supermarket with a rapid turnover of merchandise, and I do not use bleach more than three or four months old.  The change in the strength of bleach with time is yet another source of variability in the bleaching time, but this variability can be reduced by always using fresh bleach.

I have done some experiments indicating certain refinements to this bleaching protocol, and I hope to expound on them in a future update to this page.

Rinsing

Rinsing of the seed is probably not necessary as I suspect what little residual bleach remaining on the seed once the bleach is poured off is quickly dissipated.  Nevertheless I do routinely rinse the seed once the desired bleaching time is reached.  In my procedure, I pour the seed and bleach from the bottle onto a sheet of finely woven cloth cut from an old sheet or pillow case placed over the mouth of a canning jar and secured with rubber bands as shown in the figure at the right.  The cloth is pushed downward into the jar to create a depression to serve as a filter through which the bleaching solution passes but retains the seeds.  The jar and filter
Canning jar and clothare pressure sterilized along with the culture medium and are placed in a laminar flow hood.

When the bleaching of the seeds is completed, the bottle of bleach and seeds is surface sterilized by immersion in alcohol for 30 seconds and placed in the laminar flow hood.  The bottle is then opened in the hood and the contents poured into the cloth depression on top of the canning jar.  The liquid quickly drains through the cloth filter leaving the seeds deposited at the bottom of the cloth depression.  Three or four rinses of sterile reverse osmosis water are then poured over the seeds.  The entire process is done in the bath of clean air inside the laminar flow hood.  The seeds are allowed to dry for a few minutes, so that they will stick to the tip of a damp needle for placing on the medium in the culture tubes.  Many orchid labs use a procedure in which the seeds and bleach are poured onto a sheet of filter paper in a B
űchner funnel with vacuum being used to draw the bleach solution through the filter paper, but I find the procedure outlined here to be considerably more convenient.


Sowing

I use a sowing procedure that is very simple, albeit a bit tedious.  I simply prick the seeds off the cloth filter in the canning jar using a long sterile needle and place them where I want them on the surface of the planting medium.  The needles I use are homemade implements with handles.  I purchase very long needles at a craft store, and the handles are segments of plastic rod cut from a plastic coat hanger.  The blunt end of a needle is heated in the flame of an alcohol lamp and thrust into a piece of the plastic rod to create a needle with a plastic handle.


Germination Medium

In nature, terrestrial orchid seeds must be invaded by a soil fungus to germinate;  the fungus supplies nutrients and other substances necessary for germination and growth of the new protocorm.  The orchid seedling and fungus form a mycorrhizal relationship, and the fungus provides nutrients to the orchid until it reaches a stage at which it can put a green leaf above ground and obtain energy through photosynthesis.  Most Cypripedium orchids can live as autotrophs from this stage on.

While many terrestrial orchids can be germinated in the lab with the aid of a fungus, maintaining an artificial environment in which both organisms live in a balanced relationship proves difficult, and at least at the present time, germination and early growth of terrestrial orchids is carried out asymbiotically without the use of a fungus.  In asymbiotic culture, a synthetic medium, usually in the form of an agar gel, provides the orchid with all the nutrients it needs.  Years of laboratory work have gone into efforts to optimize growth media for a variety of different species.  Different species of orchids have different nutritional requirements, and individual propagators have developed their own preferred medium for each species.  Table 1 below specifies the composition of my general purpose Cyp medium in mg/L of medium, the bulk of the medium consisting of water purified by reverse osmosis (RO).  The substances in this recipe are mostly mineral nutrients required by the young orchid.  I have found that most Cyp species germinate and grow reasonably well on this medium although there are many species that do better on slight variations of this medium.  For example, C. reginae  protocorms grow much faster with 500 mg/L casein hydrolysate, whereas C. calceolus and C. kentuckiense do better with less casein.  As described below, a sugar is also essential to provide energy to the growing plants.

Medium TableIn preparing the medium, I usually weigh out the major constituents individually but add the minor and trace elements from stock solutions.  In Table 1, the constituents in the red area are contained in one stock solution, and those in the green area are in a separate stock solution.  The ingredients are initially added to roughly 0.9 L of RO water, and after all the items in the table are included, a sugar is added as described subsequently, and the pH is adjusted to the desired level by addition of 0.5 N KOH solution.  Finally the volume of the medium is brought up to 1.0 L by addition of RO water.  In actual practice, I usually add the agar after the medium has been brought up to 1.0 L volume.  As for the optimal pH of the medium, I usually choose something in the range of 6.0 to 6.3.  This is a bit higher than many orchidists use, and I like the higher pH because the plants add H+
ions to the medium during incubation resulting in a gradual drop in medium pH.  The drawback to increasing the pH to higher values is that some calcium and phosphorus precipitate out of the medium as calcium phosphate.  Thus the choice of pH is a compromise between keeping the plants in the range in which they are happy and retaining all of the calcium and phosphorus in the medium.

Three of the most important medium ingredients are not shown in Table 1, namely, a cytokinin, raw Russett potato, and a sugar.  Adding a cytokinin is not necessary for germinating seeds of many Cyp species, but is quite helpful for some including C. arietinum and C. reginae.  I have experimented with several cytokinins and found that kinetin (6-Furfurylaminopurine) and BA (6-Benzylaminopurine) are quite effective in stimulating embryo and early protocorm growth in concentrations of 0.1 to 0.5 mg/L.  While beneficial in the germination medium, cytokinins cause abnormal protocorm development, particularly as the first-root growth stage is approached, and therefore using the lowest concentration that is effective is a good plan. Moreover, replating of the protocorms from the germination medium to a medium without added cytokinin is important.  There are stronger cytokinins including meta-Topolin (6-(3-Hydroxybenzylamino)purine) and TDZ (Thidiazuron) that are even more effective in promoting embryo and early protocorm growth, but these are very deleterious to subsequent development and demand that the protocorms be replated to cytokinin-free medium at a very early stage.  I routinely use kinetin as my cytokinin of choice and always at the lowest concentration that is effective.

The second ingredient not listed in Table 1 is raw potato.  Potato is what is known as an undefined additive because the chemical composition of a potato is very complex, and no one really knows why addition of potato has a positive effect on germination and early growth.  I have long felt that such undefined additives are unscientific, and that if we really knew what we were doing in plant tissue culture, use of such additives would be unnecessary. Then one day I happened to attend a lecture by a leading cancer researcher and learned from his talk that even in such well funded cutting edge medical research, undefined additives such as "fetal bovine serum" are used in the culture of animal cells.  While I still think that if we had complete knowledge of the requirements of our young orchids including whatever combination of cytokinins, auxins, and vitamins the potato or other undefined additives provide, we would be able to grow our plants on completely defined media.

My germination cultures consist of test tubes each containing 25 mL of medium.  To each tube I add 1-cm cubes cut from a raw Russett potato before pressure sterilization.  The amount of potato per tube varies considerably from one Cyp species to another.  Some species such as C. californicum do best with only half a 1-cm potato cube per 25 mL of medium, whereas other species such as C. candidum and C. parviflorum require three 1-cm cubes per 25 mL of medium for optimum germination and growth. Interestingly, an excess of potato above the optimal level results in very high protocorm mortality.

Orchidists in general use a wide variety of undefined additives: not only potato, but things like banana or coconut water.  Early on, I chose to work with potato because I considered the composition of things like banana or coconut water to vary too much from one fruit to another owing to the stage of ripeness among other factors.  Recently, prepackaged pure coconut water has become available at health food stores and even at large supermarkets, and this product seems much more consistent than does the liquid from one supermarket coconut to another.  I have been experimenting with the commercial pure coconut water and have found that for some species, adding a small amount of coconut water in addition to the usual potato accelerates early growth.

Finally, the medium must contain a sugar as an energy source for the growing plants.  In nature, the germinating seeds would obtain their energy from their fungal host, but in axenic culture, the growth medium itself must provide the energy in the form of a carbohydrate the young orchids can utilize.  Most often this sugar is glucose or sucrose.  Either seems entirely satisfactory in supporting Cyp growth.  I have long used glucose because of its simpler composition;  I was concerned about the breakdown of sucrose into glucose and fructose in aqueous solution.  Although subsequent experiments showed that such hydrolysis does not cause a problem in Cyp cultures, I have continued to use glucose most of the time, in large part because where I live in the Upper Midwest, glucose is not expensive;  it can be purchased as "corn sugar," "grape sugar," or "dextrose" from home wine and beer brewing supply stores.  The term "dextrose" refers to the R-  enantiomer, which is the naturally occurring form of glucose.  For almost all species, I routinely use 20 g/L glucose in the germination medium, notable exceptions being C. calceolus and C. kentuckiense for which I use a bit less.

To summarize, my general germination medium has the composition:    [Table 1]  +  [20 g/L glucose (or sucrose)]  +   [0.0 to 0.5 mg/L kinetin]  +  [20 to 120 potato cubes per L]


 Incubation

After sowing the seeds on the germination medium, the cultures are placed in a box that is moved to a cabinet for incubation in the dark.  Most Cyp seeds will not germinate in the light;  they require the darkness that they would experience underground in nature.  I once found seeds of C. parviflorum var. makasin that would germinate in dim room light, but even they germinated better in darkness.  

Resist the temptation to examine cultures in the light.  Exposure to light for even a few minutes during the germination process kills germinating embryos and new protocorms of many species.  Of course, looking at the cultures to check for contamination is important, so what is one to do?  I routinely sow seeds in sets of 10 culture tubes containing 25 mL of medium, and I consider one of these a "sacrifice tube," which I examine to check for germination and contamination while maintaining the other nine tubes in darkness.  Even using such a sacrifice tubes, I still wait at least a month to examine it in the light for the first time.  Germination can usually be seen using a 10 power hand lens, but a binocular dissecting microscope magnifying 20 power or more gives a much better view.

The proper temperature during incubation is important.  Most Cyp seeds germinate well and the protocorms grow well at roughly room temperature, that is, 18 to 22 C (64 to 72 F).  There are exceptions, however;  for example seeds of C. irapeanum germinate better and the protocorms grow better at temperatures a bit above 22 C.


Replating


Replating is simply using a sterile needle to move protocorms or seedlings from the germination medium to the replating medium.  I use the same home made needles I use for sowing in replating.  

When to Replate

After successful germination and several months of incubation in the dark at room temperature, protocorms or young seedlings (An orchid seedling is a plant with a root.) become too crowded, and the medium becomes too depleted in nutrients for further growth.  I usually replate seedlings when roughly one third to one half of the protocorms have entered the first-root stage.  There are several factors that determine the optimal timeC. reginae culture tubes ready to replate for replating.  The photo at right shows C. reginae seedlings and protocorms at a good state for replating.  For some species, large protocorms and seedlings survive transplanting better than small protocorms.  In cases where the germination medium includes a cytokinin, this growth regulator may cause abnormal development as growth progresses, in which case moving the protocorms to a new medium without the cytokinin at an early stage when the protocorms are still quite small is important.  In some species, the development of root hairs known as rhizoids (visible in the three photos at the top of this page) is a practical consideration for when to replate because as the protocorms or seedlings get larger, their hairs become entangled making separation of the plantlets during replating very difficult.



Replating Medium

The replating medium usually differs from the germination medium in several important ways: 1) No cytokinin is added to the replating medium. 2) The type and quantity of gelling agent is adjusted to give a mechanically weaker gel than the germination medium, and 3) The number of potato cubes per liter of medium may be different for the replating medium.

As mentioned above, excessive cytokinin in the germination medium may interfere with normal differentiation of tissues and development of organs as the plants grow.  Some species are much more sensitive to the cytokinin than others in this regard,  but in no case is there a need for kinetin or other cytokinin to be added to the replating medium.

The relatively large amount of agar added to the germination medium produces a strong gel.  The reason so much agar is used is to reduce the amount of residual water in the culture tubes, so that the seeds to not float around randomly on the surface of the gel after sowing.  The first roots of the little plants may or may not be able to penetrate this gel, so replating to a medium with weaker gel is helpful in facilitating root growth.  When using agar for the replating medium, I usually use as little as 4.0 g/L.  Most often, however, I use a gellan gum as the gelling agent in my replating medium.  Gellan gum has several advantages over agar, one being that it makes a clearer gel than does agar.  Gellan gum also cleans from glassware much more easily than does agar.  Finally, gellan gum is cheaper to use than agar.  Kilogram for kilogram, the gum powder is more expensive than agar, but far less gum is needed to gel the medium.  I normally use only 0.3 to 0.35 g/L gellan gum, making a nice weak gel that roots can easily penetrate.   For gellan gum to form a gel, some divalent cations, particularly Ca+2
, are needed, and in special purpose replate media such as the one I use for C. arietinum, which has a low calcium concentration, a bit more gellan gum is necessary.

Flask of C. kentuckiense seedlingsAn important consideration for replating is how much medium should be allocated to each seedling.  Clearly, cramming more seedlings into a flask with a given volume of medium would be cheaper and quicker, but there is a cost in reduced seedling growth.  I normally replate seedlings at a density that gives 10 mL of medium to each seedling.  For example, I most often place 20 seedlings on 200 mL of medium in a 500 mL flask.  The photo at the left shows a 500 mL flask of C. kentuckiense seedlings.

After transfer of protocorms and seedlings to the flask, it is incubated in the dark for several additional months, the actual number depending considerably upon the species.  Some species are ready to be removed from the flask after five or six months of further growth, whereas some require considerably longer.  The total length of incubation required to produce seedlings is very helpful in deciding when to sow the seeds so as to produce seedlings ready for planting in spring. I routinely replate seedlings only once, transferring them directly from the germination medium to the final flask.  This procedure seems to produce seedlings adequate in size for planting out, but I have seen larger seedlings produced by use of a second replating to fresh medium.






















Deflasking Seedlings

When to Deflask

Several factors determine when to remove the seedlings from the flask, rinse off the gel, and place them in the refrigerator for several months of vernalization. Clearly, if the seedlings stop growing or if their root tips start turning brown, the seedlings should be removed pronto.  More often, however, the seedlings may still be growing, but yet timing dictates that they should be removed from the flask and refrigerated so as to have adequate cool time for vernalization and be planted out in the spring.  In this case, the grower must make a judgment call: Are the seedlings large and vigorous enough to produce an aerial shoot after vernalization?  One of the best clues is the size of the shoot bud;  it must be large enough to indicate sufficiently developed leaves for growth next season.  There is considerable variation of the size of the bud among species, but as a generalization, I recommend that the bud be at least 0.5 cm tall and preferably closer to 1 cm.

Upon removal from the flask, any gel clinging to the roots should be washed off under a strong spray of water.  The ideal would be to wash off every bit of the gel so as not to provide nutrients to encourage microbes or fungi to grow during refrigeration, but in species in which the orchid roots are hairy, removing all the gel is sometimes difficult.  Usually, a little remaining gel is not harmful because it has been thoroughly depleted in many nutrients by the plants.

Refrigeration

After the seedlings have been thoroughly rinsed, I usually place them in plastic food storage boxes or trays, purchased at a supermarket for refrigerating the seedlings.  Such trays are not quite airtight thus permitting gas exchange, but at refrigerator temperatures, the seedlings usually do not desiccate providing they are still wet from rinsing when placed in the box.  Depending on the size of the seedlings and the size of the tray, I usually place between 50 and 100 seedlings in each tray.  Occasionally, when I have a great number of seedlings, I may use larger trays and place as many as 200 in each.  During refrigeration, the seedlings should be checked from time to time, and if they appear to be drying out too much, add just a little water, no more than a few mLs, to the bottom of the tray.  I also vernalize small numbers of seedlings in food freezer bags.  Some propagators use similar polyethylene freezer bags even for large numbers of seedlings without any problems.  Be sure, however, to use freezer bags;  lighter weight bags permit water loss at a rate high enough that the seedlings will desiccate after several weeks.

For proper vernalization, the temperature should be held just slightly above freezing: 0 to 4 C (32 to 39 F).  The closer the temperature is to freezing, the more rapidly the seedlings vernalize, but great care must be taken to prevent freezing.  Preventing freezing in the refrigerator is often difficult because these devices usually have large temperature gradients and cold spots.  No doubt most people have had the experience that upon dialing down the temperature in the fridge to keep the milk from spoiling, the lettuce in another spot freezes.  Why freezing of Cyp seedlings just out of the flask kills them but freezing of outdoor seedlings of the same size in the ground does not, is a mystery.  Clearly somehow growing in the soil and subject to natural conditions hardens the seedlings to withstand being frozen solid.  Freezing of deflasked seedlings in the refrigerator invariably kills them.

Although the seedlings are not maintained in sterile conditions  during refrigeration, microbial infection is usually not a problem.  The only exception that I have had has been with long term refrigeration of C. reginae seedlings.  Infection of C. reginae seedlings is usually not a problem for refrigeration up to three months, but I generally refrigerate the plantlets for at least four months because in nature our northern Minnesota plants are in freezing or near-freezing temperatures for a good five months.  I have found that some, but not all, trays of C. reginae seedlings become infested with yeast over their outer surfaces.  The yeasts apparently feed on metabolic waste products exuded by the plants during dormancy.  Such yeast-infected plants eventually acquire an alcoholic smell, and their roots gradually become limp followed by death of the plant.  I have found that monthly rinsing of the seedlings with fresh water reduces the incidence of the problem, but even  with the rinsing, a small number of trays develops the yeast infection.  I am currently experimenting with rinsing the seedlings with a weak preparation of the fungicide chlorothalonil as a preventive measure.  So far I have found that the seedlings tolerate exposure to the fungicide, but I have not yet seen whether this treatment prevents yeast growth.  Fortunately, the yeast problem has not appeared with other Cyp species, not even with refrigeration up to five months.

Following refrigeration sufficient to vernalize the seedlings, they can be rinsed and planted out in a suitable soil or mix, the composition of which should be appropriate for each species.

Future Developments

I work continually to improve my lab methods, and my intention is to publish them here as time permits.   Every fall and winter I work in the lab at both sowing and replating, and I am always experimenting with tweaks to sowing and replating media for different species.  I also work at improving the pre-sowing protocol for seeds in an effort to improve both percentage germination and the speed of germination.  While I plan to update this page with results from this work, don't bother to check for monthly updates!  Yearly will be the best you can expect.

Good luck with your propagating!

Bill
August 2015

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